Tag: osteogenesis

Mesenchymal stem cells possess a cell-surface protein called ALK2. ALK2 acts as a receptor for bone-inducing growth factors. ALK2, for example, is expressed in cartilage and if mesenchymal stem cells express a constantly-active form of ALK2, known as caALK2, these cells are driven to become cartilage-making cells (known as chondrocytes).

Can this receptor be used to drive bone formation? It turns out that manipulating ALK2 can drive fat-based stem cells (ASCs) to become bone making cells that ultimately improve bone tissue engineering. Researchers from the laboratory of Benjamin Levi at Massachusetts General Hospital, Boston, Massachusetts have fiddled with ALK2 in mesenchymal stem cells to for formation of bone from ASCs, and to enhance bone regeneration in a living animal.

To do this, Levi’s team genetically manipulated mice so that they expressed a form of ALK2 that was constantly turned on known as caALK2. The fat-based MSCs were then isolated and analyzed for their ability to make bone in culture. caALK ASCs were much more responsive to bone-inducing growth factors. These cells also expressed a whole host of bone-specific genes (e.g., Alp, Runx2, Ocn, Ops) after seven days. Since the caALK2 MSCs did so well in culture, they were then tested in mice with skull defects. Bone formation was significantly higher in mice treated with caALK2-expressing ASCs than those treated with normal ASCs.

Thus, Levi’s laboratory has shown that by treating mice with fat-based stem cells that express a constitutively active ALK2 receptor showed significantly increased bone formation. This increased bone formation can also be harnessed to improve skull healing in mice with bone defects.

Repairing cartilage defects in the knee represents one of the primary goals of orthopedic regenerative medicine. Cartilage that covers the joints, otherwise known as articular cartilage, has a limited capacity for repair, which leads to further degeneration of the cartilage when it is damaged if it remains untreated. A number of surgical options for treating cartilage defects include microfracture, osteochondral grafting, and cell-based techniques such as autologous chondrocyte implantation (ACI). Each of these procedures have been used in clinical settings. Unfortunately cartilage injuries treated with microfracturing deteriorate with time, since the cartilage made by microfracturing has a high proportion of softer. less durable fibrocartilage. Also osteochondral grafting suffers from a lack of lateral integration between host and donor cartilage.

Alternatively, tissue engineering has shown some promise when it comes to the healing of cartilage defects. Mesenchymal stem cells (MSCs) are multipotent progenitor cells that have the ability to differentiate into several different cell lineages including cartilage-making chondrocytes. MSCs have theoretical advantages over implanted chondrocytes when it come to healing potential. MSCs have the ability to proliferate without losing their ability to differentiate into mature chondrocytes and produce collagen II and aggrecan. In the short-term, bone marrow-derived MSCs combined with scaffolds have been successful in cartilage repair using animal models such as rabbits (Dashtdar H, et al., J Orthop Res 2011; 29: 1336-42) sheep (Zscharnack M, et al., Am J Sports Med2010; 38: 1857–69) and horses (Wilke MM, et al.,l J Orthop Res2007; 25: 913–2).

In a recent study, Kazumasa Ogasawara and Yoshitaka Matsusue and their colleagues from Shiga University of Medical Science in Shiga, Japan, tested the ability of expanded bone marrow-derived MSCs that had been placed in a collagen scaffold to improve healing of cartilage defects in cynomolgus macaques (type of monkey). Before this study, there were no previous studies using MSCs from primates for cartilage repair. The monkey MSCs were shown to properly differentiate into fat, bone, or cartilage in culture, and then were transplanted into the injured cartilage in the cynomolgus macaque. The efficacy of these cells were ascertained at 6, 12, and 24 weeks after transplantation.

In culture, the cynomolgus MSCs were able to differentiate into fat, bone, and cartilage.

Upon transplantation into cartilage defects in the knee cartilage of cynomolgus monkeys, MSCs were compared with collagen gel devoid of MSCs. The knees that received the transplantations did not show any signs of irritation, bone spurs or infection. All of the animals had so-called “full-thickness cartilage defects,” and those in the non-treated group showed cartilage defects that did not change all that much. The cartilage defects of the gel group had sharp edges at 6 weeks that were thinly covered with reparative tissue by 12 weeks, and at 24 weeks, the defect was covered with thick tissue, but the central region of the defects often remained uncovered, with a hollow-like deformity. In the cartilage defects of those animals treated with MSCs plus the collagen gel, the sharp edges of the defects were visible at 6 weeks after the operation, but at 12 weeks, the defects were evenly covered with yellowish reparative tissue. At 24 weeks, the defects were covered with watery hyaline cartilage-like tissue that was very similar to the neighboring naïve cartilage.

Macroscopic observations of the repaired defects in the 3 groups at 6 weeks (a, d, g), 12 weeks (b, e, h), and 24 weeks (c, f, i) after implantation. Scale bar: 5 mm. Arrow in (d): the sharp edge of the defect is visible at 6 weeks in the gel group. Arrow in (f): a hollow-like deformity remains in the central region of the defect, despite thick coverage by the reparative tissue. Arrow in (g): the sharp edge of the defect is also visible in the MSC group at 6 weeks.Read More: http://informahealthcare.com/doi/full/10.3109/17453674.2014.958807.

When evaluated at the tissue level, Ogasawara and Matsusue and others used a stain called toluidine blue to visualize the amount of cartilage made by each treatment. As you can see in the picture below, the non-treated group didn’t do so well. In the full-thickness defect the region below the cartilage was filled with amorphous stuff 6 weeks after the procedure, and at 12 weeks, amorphous stuff faintly stained with toluidine blue, which reflects the conversion of the amorphous stuff into bone. At 24 weeks, bone tissue reappeared below the cartilage zone, even though the bone did not look all that normal (no trabecular structure but woven bone-like structure).

In the gel group, cartilage-like tissue is seen at 6 weeks, and at 12 weeks, the faintly stained layer covered the cartilage defect. At 24 weeks, the defect was covered with the cartilage-like stuff, even though the central region had only a little cartilage, as ascertained by toluidine blue staining. The bone underneath the cartilage looked crummy and there was excessive growth of cartilage into the region underneath the cartilage layer.

In the MSC group, the bone underneath the cartilage healed normally, and at 12 weeks, the boundary between the articular cartilage and the bone layer beneath it had reappeared. At 24 weeks, the thickness of the toluidine blue-stained cartilage layer was comparable to that of the neighboring naïve cartilage.

Even though the gel group showed most cartilage-rich tissue covering the defect, this was due to the formation of excessive cartilage extruding through the abnormal lower bone layer. Despite the lower amount of new cartilage produced, the MSC group showed better-quality cartilage with a regular surface, seamless integration with neighboring naïve cartilage, and reconstruction of the bone underneath the cartilage layer.

This protocol has been nicely optimized by Ogasawara and Matsusue and their research team. From these data, they conclude: “Application in larger defects is certainly in line with future clinical use. If MSCs—under optimized conditions—turn out to be superior to chondrocyte implantation in experimental cartilage repair, the procedure should be introduced to clinical practice after well-controlled randomized clinical trials.” Hopefully, clinical trials will commence before long. This procedure uses a patient’s own MSCs, and if such a procedure could reduce or delay the number of knee replacements, then it would surely be a godsend to clinicians and patients alike.

Johnny Huard and his co-workers from the McGowan Institute for Regenerative Medicine at the University of Pittsburgh have isolated a slowly-adherent stem cell population from skeletal muscle called muscle-derived stem cells or MDSCs (see Deasy et al Blood Cells Mol Dis 2001 27: 924-933). These stem cells can form bone and cartilage tissue in culture when induced properly, but more importantly when MDSCs are engineered to express the growth factor Bone Morphogen Protein-2 (BMP-2), they make better bone and do a better job of healing bone lesions than other engineered muscle-derived cells (Gates et al., J Am Acad Orthop Surg 2008 16: 68-76).

In most experiments, MDSCs are infected with genetically engineered viruses to deliver the BMP-2 genes, but the use of viruses is not preferred if such a technique is to come to the clinic. Viruses elicit and immune response and can also introduce mutations into stem cells. Therefore a new way to introduce BMP-2 into stem cells is preferable.

To that end, Huard and his colleagues devised an ingenious technique to feed BMP-2 to implanted MDSCs without using viruses. They utilized a particle composed of heparin (a component of blood vessels) and a synthetic molecule called poly(ethylene arginylaspartate diglyceride), which is mercifully abbreviated PEAD. The PEAD-heparin delivery system formed a so-called “coacervate,” which is a tiny spherical droplet that is held together by internal forces and composed of organic molecules. These PEAD-heparin coacervates could be loaded with BMP-2 protein and they released slowly and steadily to provide the proper stimulus to the MDSCs to form bone.

When tested in culture dishes, the BMP-2-loaded coacervates more than tripled the amount of bone made by the MDSCs, but when they were implanted in living rodents the presence of the BMP-2-loaded coacervates quadrupled the amount of bone made by the MDSCs.

This technique provides a way to continuously deliver BMP-2 to MDSCs without using viral vectors to infect them. These carriers do inhibit the growth or function of the MDSCs and activate their production of bone.

This paper used a “heterotropic bone formation assay” which is to say that cells were injected into the middle of muscle and they formed ectopic bone. The real test is to see if these cells can repair actual bone lesions with this system.

Researchers from Boston, MA have used synthetic silicate nanoplatelest or layered clay to induce bone cell formation from stem cells in the absence of other bone-inducing factors.

Synthetic silicates are composed of either simple or complex salts of silicic acid (SiH4O4). Silicic acids have been used extensively in commercial and industrial applications that include food additives, glass and ceramic filler materials, and anti-caking agents.

In this study, novel silicate nanoplatelets were constructed that stimulated human mesenchymal stem cells to differentiate into bone-making cells in the absence of any bone-inducing growth factors or cytokines. The presence of the silicate triggers a set of events inside the mesenchymal stem cells that re-enacts the steps cell normally take during development when they form become bone cells. These exciting findings illustrate how the use of these silicate nanoplatelets in designing bioactive scaffolds for tissue engineering can lead to the formation of clinically useful bone tissues.

The lead author of this work, Ali Khademhosseini from the division of biomedical engineering at Brigham and Woman’s Hospital, thinks that silicic acid derivatives might be useful in engineering bone. “With an aging population in the U.S., injuries and degenerative conditions are subsequently on the rise,” said Khademhosseini. This means that there is also an increased demand for therapies to repair damaged tissues. Forming such tissues requires protocols to direct stem cell differentiation so that the cells can form new tissues and biomaterials. According to Khademhosseini, “Silicate nanoplatelets have the potential to address this need in medicine and biotechnology.”

“Based on the strong preliminary studies, we believe that these highly bioactive nanoplatelets may be utilized to develop devices such as injectable tissue repair matrixes, bioactive filters, or therapeutic agents for stimulating specific cellular responses in bone-related tissue engineering,” said Akhilesh Gaharwar, first author of this present study.

Future mechanistic studies are necessary to elucidate those underlying pathways that govern the induction of bone differentiation by materials like silicates. Such studies should lead to a better understanding of how particular strategies can be adjusted to improve the performance of lab constructed biomaterials, and accelerate patient recovery time.

Patient-specific bone substitutes have been produced by a team of scientists from the New York Stem Cell Foundation. Darja Marolt and Giuseppe Maria de Peppo from the New York Stem Cell Foundation (NYSCF) led the study that demonstrated that customizable, three-dimensional bone grafts that can be produced on-demand for patients from their own cells.

Marolt and de Peppo and their co-workers used skin grafts from their patients to isolate skin fibroblasts that were reprogrammed to induced pluripotent stem cells (iPSCs). Because iPSCs are made from the patient’s own cells, they have the same profile of cell surface proteins as the patient’s own tissues. Therefore, they are very unlikely to be rejected by the patient’s immune system. Also, iPSCs have the ability to differentiate into any cell type found in the adult body, and therefore, can be used to form bone cells.

After deriving iPSCs from patient skin cells, de Peppo, Marolt, and colleagues coaxed the cells to form osteoblasts (the cells that form bone), and seeded them onto a scaffold that mimicked three-dimensional bone structure. These structures were grown in a bioreactor that fed the cells oxygen and nutrients.

According to Marolt, “Bone is more than a hard mineral composite, it is an active organ that constantly remodels. Blood vessels shuttle important nutrients to healthy cells and remove waste; nerves provide connection to the brain; and, bone marrow cells form new blood and immune cells.”

Previous studies have demonstrated that cells from other sources also possess bone-forming potential. However, these same studies have revealed serious shortcomings of the clinical potential of such cells. A patient’s own bone marrow stem cells can form bone and cartilaginous tissue, but not the accompanying underlying vasculature and nerve compartments. Also, embryonic stem cell derived bone may prompt an immune rejection. Therefore, the use of iPSCs can overcome many of these limitations.

As de Peppo noted: “No other research group has published work on creating fully viable functional three-dimensional bone substitutes from humans iPS cells. These results bring us closer to achieving our ultimate goal, to develop the most promising treatments.”

Since bone injuries and defects are often treated with bone grafts that are taken from other parts of the body or a tissue bank. Alternatively, synthetic alternatives can also be used, but none of these possibilities provide the means for complex reconstruction and they may also be rejected by the immune system, or fail to integrate with surrounding connective tissue. n the case of trauma patients who suffer from shrapnel wounds or vehicular injuries , the traditional treatments provide only limited functional and cosmetic improvements.

To access the integrity of the bioreactor-made bone, the NYSCF team implanted them into animals. Implantation of undifferentiated iPSCs formed tumors, but transplantation of the iPSC-derived bone produced no tumors, but also produced grafts that effectively integrated into the bones, connective tissues and blood vessels of the animals.

Susan Solomon, CEO of NYSCF, said of this work, “Following from these findings, we will be able to create tailored bone grafts, on demand, for patients without any immune rejection issues. She continued: “it is the best approach to repair devastating damage or defects.”

The therapeutic relevance of this work aside, these adaptive bone substitutes can also serve as models for bone development and various bone pathologies. Such bone exemplars could serve as models for drug testing and drug development.

If MSCs throughout the body are similar cell types then we would expect them to have similar embryological origins. However, this is not the case, since MSCs develop from several different embryonic tissues. The first wave of MSCs arises from Sox-1-expressing neuroepithelial cells during embryonic development. However, later MSCs come from multiple sources (Takashima et al 2007), including neural crest cells (Nagoshi et al 2008; Morikawaet al 2009). Therefore, MSCs from various tissues almost certainly have distinct embryological origins. Additionally, MSCs are located in different sites in the body, and are influenced by specific microenvironments. Thus MSCs from different tissue sources might represent distinct cell types, and could potentially display distinct differentiation profiles and express particular genes. Despite these differences in developmental origin and environmental influences, MSCs from various sources have very similar morphologies and share a common array of surface markers (Mitchell et al 2003; Lee et al 2004a; Wang et al 2004; Tsai et al 2007). However, several studies have established that MSC populations are rather heterogeneous (Dominici et al 2009), and, therefore, surface markers expressed on some cells of an MSC population are not always expressed in all the cells of that population (Mafi et al 2011). Also, the growth kinetics of cultured MSCs differs remarkably with respect to their source (Kang et al 2004b; Yoshimura et al 2007; Troyer and Weiss 2008).

Despite the shared array of cell surface markers, presently there are no cellular markers or cell surface proteins that are unique to MSCs. In order to provide a more unified approach to MSC biology, the International Society of Cryotherapy has proposed three criteria for the identification of MSCs. Under these criteria, MSCs must: (1) be plastic-adherent when maintained in standard culture conditions; (2) express the following cell surface molecules CD105, CD73 and CD90, and lack expression of CD45, CD34, CD14 or CD11b, CD79alpha or CD19 and HLA-DR surface molecules, and; (3) be able to differentiate into osteoblasts, adipocytes and chondroblasts in vitro (Dominici et al 2006). Despite these definitions, flow cytometric analyses of MSCs from several different populations have shown some significant differences in cell surface markers (Boeuf and Richter 2010). For example, even though the absence of CD34 is generally considered a criterion for the definition of MSCs, various investigators have reported low expression of CD34 in ADSCs (ADSCs; De Ugarte et al 2003a; Rebelatto et al 2008; Roche et al 2009) and BMSCs (Zvaifler et al 2000; Gronthos et al 2003; Yu et al 2010). Likewise, many investigators have shown that MSCs from multiple sources do not express CD45 (Zvaifler et al 2000; Zuk et al 2002; Igura et al 2004; Dominici et al 2006; Wongchuensoontorn et al 2009), but BMSCs are CD45 positive (Yu et al 2010).

Comparative gene array analyses of MSCs from different sources have revealed some differences in gene expression between these distinct MSC populations, but overall the gene expression profiles between these cells are relatively similar (Winter et al 2003; Lee et al 2004a; Djouad et al 2005; Wagner et al 2005; Aranda et al 2009; Jansen et al 2010). Proteomic comparisons of distinct MSC populations using two-dimensional gel electrophoresis analysis came to very similar conclusions (Roche et al 2009). MSCs from intra-articular tissues (synovial membrane and anterior cruciate ligament) and chondrocytes show gene expression profiles that were more similar to each other than to MSCs from extra-articular locations (Segawa et al 2009). These data suggest that MSCs from varied sources probably represent similar, but distinct cell types that express a core of common genes, but also clusters of distinct genes. These gene expression differences convey different differentiation potentials upon specific MSC populations and varied requirements for these particular MSC populations to differentiate into specific cell types (Gimble et al 2008; Rastegar et al 2010).

MSC Differentiation
With respect to the differentiation potential of MSC populations, the general rule of thumb is the closer the MSC source tissue is to the target tissue, the more effectively that particular MSC population differentiates into the target tissue. A few examples should suffice. Yoshimura and colleagues found that rat SMSCs derived from the synovial tissue of the knee, which is closest to the target tissue of chondral cartilage, formed cartilage better than BMSCs, ADSCs, or MSCs from periosteum or muscle (Yoshimura et al 2007). Likewise, gene expression profiles of human BMSCs or umbilical cord-derived MSCs (UCSCs from Wharton’s jelly) definitively showed that BMSCs express a variety of osteogenic genes (RUNX2, DLX5 and NPR3) not observed in UCSCs. Under osteogenic induction, BMSCs produced far more bone than UCSCs. However, UCSCs express angiogenesis genesand fewer genes involved in the immune response than BMSCs, suggesting that UCSCs are superior for allogeneic transplantation. When cocultured with allogeneic macrophages,UCSCs prevented the macrophages from producing immunomodulatory cytokines tumor necrosis factor and Interleukin-6 (Hsieh et al 2010). Finally, Niemeyer and coworkers showed that BMSCs and ADSCs formed bone with similar efficiencies in vivo (Niemeyer et al 2007), but in animals studies, BMSCs produced better repair of tibial osteochondral defects in sheep when compared to ADSCs (Niemeyeret al 2010).

MSC Chondrogenesis
Initiation of cartilage development during animal development begins with the condensation of mesenchymal precursor cells (Woods, Wang and Beier 2007). These cell-cell contacts are mediated by N-cadherin, whose expression is highly upregulated in human MSCs after being subjected to chondrogenic induction (Tuli et al 2003). N-cadherin is required for chondrogenesis of chick limb mesenchymal cells in vitro and in vivo (Oberlender and Tuan 1994). Prior to MSC condensation prechondrocytic MSCs secrete extracellular matrix rich in hyaluronic acid, collagen type I and IIa. Initiation of MSC condensation also correlates with the expression of neural cadherin (N-cadherin) and neural cell adhesion molecule (N-CAM). The secreted signaling molecule transforming growth factor-β (TGF-β) is one of the earliest signals in chondrogenic condensation. TGF-β activates production of the extracellular matrix protein fibronectin, which up-regulates N-CAM, and also stimulates the synthesis of Sox transcription factors (Sox-5, -6 and -9), which are essential for cartilage formation. Other extracellular matrix molecules made by chondrogenic MSCs include tenascins, thrombospondins, and cartilage oligomeric protein (COMP). These extracellular matrix molecules interact with cell adhesion molecules to activate intracellular signaling pathways that initiate the transition from chondroprogenitor cells to fully committed chondrocytes. Proliferating chondroprogenitor cells synthesize hyaluronan, collagen II, IX and XI, and the cartilage-specific proteoglycan core protein (or chondroitin sulfate proteoglycan 1) known as aggrecan. Aggrecan (encoded by the ACANgene) is a member of the aggrecan/versican proteoglycan family, and is the most predominant proteoglycan in the extracellular matrix of articular cartilage. Aggrecan helps cartilage withstand compression. N-cadherin and N-CAM expression fade and disappear in differentiating chondrocytes (Golding, Tsuchimochi and Ijiri 2006).

When grown under chondrogenic conditions, MSCs in monolayer culture respond by condensing into high-density three-dimensional cell aggregates (Winter et al 2003). In order to realistically recapitulate chondrogenesis in culture, researchers deposit centrifuged MSC pellets that contain ~200,000 – 500,000 cells in a two-dimensional culture. This culture system, which is one of the most widely used in chondrogenesis research, is called a pellet, aggregate or spheroid culture. To induce chondrogenesis, pellets are cultured in a basal medium (typically low- or high-glucose Dulbecco’s Modified Eagles Medium, otherwise known as DMEM, or fetal calf serum) that contains dexamethasone, ascorbate, proline, insulin, transferrin and selenous acid (Johnstone et al 1998; MacKay et al 1998; Puetzer, Petitte and Loboa 2010). Classically, the growth factor used to induce chondrogenesis in this type of medium is 10 ng/ml of transforming growth factor-β (TGF-β). TGF-β1, 2, and 3 are the only well-established full inducers of chondrogenesis that, when added as single factors, induce proteoglycan and collagen type II deposition (MacKay et al 1998; Barry et al 2001). Other chondrogenic inducers have been described; bone morphogen protein-2 (BMP-2) for BMSCs (Schmitt et al 2003) and BMP-6 for ADSCs (Estes, Wu and Guilak 2006). However, other studies have failed to confirm the chondrogenic efficacy of these two growth factors (Winter et al 2003; Indrawattana et al 2004; Xu et al 2006; Hennig et al 2007; Weiss et al 2010), and there is even a chance that these two growth factors might only work in a donor-specific fashion.BMP-2, -4, and -6, and insulin-like growth factor-1 (IGF-1) seem to promote chondrogenesis in MSCs when given in combination with TGF-β (Schmitt et al 2003; Im, Shin and Lee 2005; Sekiya et al 2005; Liu et al 2007).

These data do not necessarily mean that BMSCs are the best cartilage-making MSCs in the body. First of all, head-to-head comparisons treated both MSC populations with the same chondrogenic induction protocol, which implicitly assumes that culture conditions optimized for BMSCs are also be optimal for ADSCs. This assumption, however, ignores the intrinsic differences between these two MSC populations. Kim and Im have shown that ADSCs display a chondrogenic potential equal to that of BMSCs if ADSCs are treated with higher concentrations of growth factors (Kim and Im 2009). Additionally, Diekman and colleagues have shown that chondrogenesis of BMSCs and ADSCs is highly dependent on the presence and concentration of particular growth factors, the presence or absence of serum, and the composition of the scaffold in which the cells are embedded for the chondrogenic induction. ADSCs made significantly more aggrecan in response to BMP-6 than to TGF-β, but the opposite was true for BMSCs. Likewise, ADSCs produced more type II collagen in the presence of serum whereas BMSCs produced more type II collagen without serum. Finally when seeded in alginate beads, the quantity of glycosaminoglycan (GAG) made by BMSCs were significantly higher in the dual-growth factor cocktail of TGF-β and BMP-6 as compared to TGF-β alone. However, when these same cells were grown in a cartilage-derived matrix, those grown in the TGF-β-alone cocktail had higher viability and produced higher amounts of GAG when compared to those grown in dual cocktail (TGF-β + BMP-6). Thus the growth scaffold greatly influences the response of MSCs to particular growth factors, but these data also underscore that BMSCs and ADSCs are probably distinct cell types (Diekman et al 2010).

Secondly, keeping with the original rule that the closer the source tissue is to the desired target tissue, the more effectively MSCs from those tissue sources differentiate into the target tissue, Sakaguchi and colleagues showed that MSCS from bone marrow, synovium, and periosteum made more cartilage than ADSCs or skeletal muscle-derived MSCs, but SMSCs clearly made the most cartilage (Sakaguchi et al 2005). Interestingly, this result was replicated in rat MSCs (Yoshimura et al 2007). Equine BMSCs, however, do show superior chondrogenesis to UCSCs and MSCs from amniotic fluid (Lovati et al 2011), and human fetal and adult BMSCs exceed the chondrogenic potentials of fetal lung-, and placenta-derived MSCs (Bernardo et al 2007).

The varied responses of MSCs from various sources to different growth factors also have been well documented. For example, TGF-β alone is sufficient for chondrogenesis of BMSCs (Afizah et al 2007), but not ADSCs (Awad et al 2003: Estes, Wu and Guilak 2005). Additionally, the combination of TGF-β and dexamethasone stimulates chondrogenesis in BMSCs, but in ADSCs, TGFβ is required for chondrogenesis but dexamethasone tends to suppress chondrogenesis (Awad et al 2003). The reduced chondrogenic induction of ADSCs by TGF-β is probably due to reduced expression of the TGF-β receptor in these cells. However, BMP-6 treatment induces expression of the TGF-β receptor ALK-5 in ADSCs and combined application of TGF-β and BMP-6 restores chondrogenesis in this MSC population (Hennig et al 2007). A published protocol to successfully differentiate ADSCs into chondrocytes makes use of the combination of TGF-β and BMP-6 (Estes et al 2010).

Differential responses to BMP-6 are also observed in different types of MSCs. As previously mentioned, BMP-6 strongly induces chondrogenesis in ADSCs, but not in BMSCs. BMP-6 in combination with TGF-β inhibits hypertrophy in ADSCs (Estes, Wu and Guilak 2003), but in BMSCs, BMP-6 promotes hypertrophy and endochondral ossification (Sekiya, Colter and Prockop 2001; Sekiya et al 2002; Indrawattana et al 2004).

These varied responses to growth factors by distinct MSC populations might also be a reflection of the assorted levels of “stemness” found among the cells of each MSC population. As previously noted, MSC populations tend to be highly heterotropic, and clonal analyses of ADSCs have shown that these cell populations are a mixture of cells that can form bone, cartilage and fat (tripotent), those that can only form two of these tissues (bipotent), and others that can only form only cell type (monopotent). The ratios of these tripotent, bipotent to monopotent clones seems to vary from study to study. Guilak and colleagues found that 21% of ADSCs clones were tripotent and approximately 30% were bipotent (Guilak et al 2006), but Zuk and others found that only 1.4% of all ADSC clones were tripotent (Zuk et al 2002). The disparities between these studies seem to be due to the media conditions used, the age of the adipose tissue donors, and the overall design of the experiment. However, these studies certainly show that distinct MSC populations consist of cells at varying levels of “stemness,” with some being more committed to a particular cell type and others being less developmentally committed to a particular cell fate. The heterogeneity of these populations almost certainly influences the response of these cell populations to particular growth factors.

In head-to-head comparisons with other types of MSCs, the osteogenic potential of BMSCs was approximately the same as SMSCs, and only slightly better than periosteum-derived MSCs (Sakaguchi et al 2005). However, in another study SMSCs from healthy donors expressed significantly lower levels of osteogenic markers after induction of osteogenesis (Djouad et al 2005). Another comparison between human umbilical cord perivascular cells (HUCPVCs) and BMSCs found that HUCPVCs had higher osteogenic potential than BMSCs (Baksh, Yao and Tuan 2007). However, other studies compared the gene expression profiles and osteogenic potential of UCSCs and BMSCs not only showed a pronounced expression of osteogenic genes in BMSCs, but also established their superior osteogenic potential in in vitro differentiation assays (Hsieh et al 2010; Majore et al 2011). It is unclear if these two experiments analyzed the same umbilical cord cell populations. MSCs isolated from human umbilical cord blood also showed a distinctly greater osteogenic potential in comparison to BMSCs (Chang et al 2006a). Also human UCSCs show superior osteogenic potential in comparison to chorionic plate-derived MSCs (Kim et al 2011).

Adipogenic induction of cultured MSCs requires the use of compounds that increase intracellular levels of the signaling molecule 3’,5’-cyclic adenosine monophosphate (cAMP) such as phosphodiesterase inhibitors (e.g., isobutylmethylxanthine or theophylline), and ligands for the glucocorticoid receptor (e.g., dexamethasone), and PPAR-γ, (i.e., rosiglitazone, which is marketed as the anti-diabetic insulin sensitizer AvandiaTM). Additionally, most adipogenic cocktails also include insulin, and some protocols also include indomethacine (Mosna, Sensebe and Krampera 2010). MSCs exposed to these agents form intracellular droplets composed of neutral lipid and express key adipogenic markers (e.g., adiponectin, fatty acid binding protein, aP2) within three-to-nine days (Gimble et al 2008; Muruganandan, Roman and Sinal 2009).

Exposing MSCs to low serum concentrations or horse serum leads to the expression of skeletal muscle markers such as myogenin and the formation of multi-nuclear myotubes. However, MSCs do not differentiate into mature, skeletal muscles as readily as they do smooth muscles, and the culture conditions under which the cells are grown seem to be extremely important. Co-culturing BMSCs (Lee, Kosinski and Kemp 2005; Beier et al 2011) or ADSCs (Di Rocco et al 2006) with skeletal muscles can induce myotube formation and the expression of myogenic genes by MSCs. The efficiency of skeletal muscle formation with this procedure is almost doubled by exposing MSCs to the chromatin remodeling reagent trichostatin A (Collins-Hooper et al; 2011). Incubation of MSCs with conditioned medium prepared from chemically damaged, but not undamaged, muscle cells also induces MSC myotube formation and expression of MyoD (Santa Maria, Rojas and Minguell; 2004). Treatment of MSCs with particular molecules such as Galectin-1 (Chan et al 2006), TWEAK (Gigenrath et al 2006) and 5-azacytidine (Kocaefe et al 2010; Natasuke et al 2010) can also induce myogenesis, as can hypoxic preconditioning (Leroux et al 2010).

Dezawa and colleagues have published a protocol for differentiating BMSCs into skeletal muscle. They treated mouse BMSCs for three days with a mixture of bFGF, forskolin, which is known to increase intracellular concentrations of cAMP, platelet-derived growth factor and neuregulin. After the three-day culture period, they transfected the cells with a plasmid that encoded the intracellular domain of the Notch receptor, and selected only those cells that had successfully taken up the plasmid. To augment the ability of the remaining cells to form myotubes, they exposed the cells to either 2% horse serum or ITS (insulin-transferrin-selenite) in serum-free medium. Both of these media promoted myogenic differentiation of MSCs to myoblasts that formed myotubes, and were able to integrate into existing muscle and repair muscle in mdx mice (Dezawa et al 2005). mdx Mice harbor a loss-of-function mutation in the gene that encodes the dystrophin protein, which, in humans, is defective in individuals who are afflicted with Duchenne Muscular Dystrophy (Muntoni, Torelli and Ferlini 2003). Therefore, even though it shows a relatively mild phenotype, the mdx mouse is a model system for muscular dystrophy (Sicinski et al 1998).

Treatment of MSCs with a drug called 5-azacytidine directs them to transdifferentiate into cells that resemble cardiomyocytes (heart muscle cells). In cells, 5-azacytidine is incorporated into DNA where it inhibits DNA methylation, and DNA hypomethylation leads to activation of particular genes (Christman 2002). Treatment of BMSCs (Fukuda 2001; Shim et al 2004; Xu et al 2004; Antonitsis et al 2007; 2008), ADSCs (Rangappa et al 2003b; Lee et al 2009) or UCSCs (Cheng et al 2003) with 5-azacytidine drives them to form cells that have a fibroblast-like morphology, synchronously beat, and express many cardiac-specific genes like troponin T, atrial natriuretic protein (ANP), GATA-4, Nkx2.5, TEF-1, and MEF-2C (Fukuda 2001; 2002; Yang et al 2012). Some work has even shown that these differentiated MSCs respond to adrenergic and muscarinic stimulation (Fukuda 2002), and can integrate into the heart of a laboratory animal and form functional connections with native cardiomyocytes (Hattan et al 2005).

Because MSC populations tend to form smooth muscle rather readily, there have been few head-to-head comparisons of the efficiency of smooth muscle formation in distinct MSC populations.

Comparisons of the ability of various MSC populations to differentiate into skeletal muscles include in vitro differentiation of MSCs from bone marrow, spleen, thymus, and liver. This study showed that BMSCs, liver- and thymus-derived MSCs all made skeletal muscle in culture, but splenic-derived MSCs did not (Gornostaeva, Rzhaninova and Gol’dstein 2006). Comparisons of the in vivo ability of BMSCs, ADSCs, and SMSCs to form skeletal muscle when implanted showed that ADSCs had the greatest ability to integrate into existing muscles (de la Garza-Rodea et al 2011).

Interestingly, a small fraction of BMSCs can form myotubes and integrate into existing muscle when injected into laboratory animals, whether that muscle is damaged or not (Ferrari et al 1998), a characteristic also shared by SMSCs (De Bari et al 2003). However, when UCSCs were injected into the tail vein of mdx mice, the cells were able to integrate into the muscle but unable to differentiate in vivo into mature, skeletal muscles (Vieira et al 2010; Zucconi et al 2011). Different MSCs show varying efficiencies of cardiomyocyte differentiation. UCSCs, for example, show particularly low transdifferentiation rates (Martin-Rendon et al 2008). ADSCs, however, transdifferentiate into cardiomyocytes with the highest efficiency (Zhu et al 2008; Tobita, Orbay and Mizuno 2011; Paul et al 2011;Yong et al 2012). In fact, when grown in a semisolid methycellulose medium enriched with growth factors, ADSCs spontaneously form beating ventricular- and atrial-like cardiomyocytes (Planat-Benard et al 2004). This makes ADSCs an attractive source of material for cardiac regenerative therapies.

MSCs and Tooth Formation
Tooth formation results from a complex set of interactions between the overlying stomadial epithelium and underlying mesenchymal cells. Dental mesenchymal cells develop from neural crest cells derived from midbrain and hindbrain cranial neural crest cells. In mice, these two cell populations are in place by day 8.5 (E8.5) and by day 10.5 (E10.5) tooth-forming sites and tooth types are determined. At E11.5, a localized thickening of the dental epithelium that results from cell shape changes forms the “dental placode.” Between E12.5-E13.5, the dental placode proliferates and invaginates to form the epithelial bud around which mesenchymal cells condense (Peters and Bailing 1999). At E14.5, the cap stage, the epithelial component of the developing tooth folds and forms a transient cluster of non-dividing cells called the “enamel knot.” The enamel knot is a signaling center that produces many powerful growth factors, including Sonic hedgehog (Shh), BMP-2, BMP-4, BMP-7, FGF-4 and FGF-9 (Thesleff and Mikkola 2002). The cap stage is followed by the bell stage, and at this time the epithelially-derived ameloblasts and the mesenchymally-derived odontoblasts differentiate. The ameloblasts form enamel and the odontoblasts produce the dentine. MSCs also generate the alveolar bone that forms the sockets for the teeth. Human tooth development occurs in a very similar fashion (Zhang et al 2005).

In adult animals, dentinal repair results from odontoblasts that differentiate from a precursor cell population that resides in dental pulp tissue. These dental pulp stem cells (DPSCs) have been isolated from adult human teeth (Gronthos et al 2002). In culture, DPSCs show robust growth and a high proliferation rate and, even after extensive subculturing, have the ability to form a dentin/mineralized complex with a mineralized matrix when grafted into the dorsal surface of immunocompromised mice (Gronthos, et al 2002; Batouli et al 2003). In a rabbit model of tooth regeneration, DPSCs are able to support the formation of functional teeth (Hung et al 2011), and in mouse and dog models, DPSCs regenerated alveolar tooth socket bone in the jaw (Yamada et al 2010; 2011; Ito et al 2011).

In a head-to-head comparison of the ability of DPSCs and ADSCs to replace teeth in a rabbit model, the teeth produced by ADSCs were very similar to those generated by DPSCs. Both sets of replacement teeth were living teeth with nerves and vascular systems, but the ADSCs grew at faster rate and were more resistant to senescence (Hung et al 2011). BMSCs, like DPSCs, are also able to form calcified deposits in vitro (Gronthos et al 2000). Likewise, gene microarray analyses of these two stem cell populations show similar levels of expression for more than 4000 genes, with only a few differences (Shi, Robey and Gronthos 2001). Head-to-head comparisons of BMSCs, DPSCs, and SHEDs have shown that these stem cells have an equivalent the ability to regenerate alveolar tooth socket bone in the jaws of laboratory animals (Yamada et al 2010; 2011; Ito et al 2011). Comparison of BMSCs and SHED gene expression profiles by means of DNA microarray and real-time reverse transcriptase polymerase chain reaction has shown that 2753 genes in SHEDs show a more than two-fold difference in expression level in comparison to BMSCs. The genes that show the greatest differences in expression in SHEDs are those involved in BMP signaling, and the protein kinase A (PKA), c-Jun-N-terminal kinase (JNK), and apoptosis signaling-regulating kinase-1 (ASK-1) signaling cascades. Therefore SHEDs have specific characteristics that differ from BMSCs, and the osteogenic and odontogenic differentiation of SHEDs and BMSCs are probably regulated by different mechanisms (Hara et al 2009).

BMSCs can probably serve as a source for dental regenerative treatments, but the faster growth rates and easier isolation of ADSCs probably makes them a superior choice.

Despite reports that MSCs can be differentiated into functional neurons, several studies have failed to recapitulate these results (Scuteri et al 2010). Time-lapse photography of rat BMSCs that had undergone neural induction showed that instead of extending neurites, the cells merely shrunk and retracted their cell extensions so that only two extensions remained. This was interpreted to be a response to toxic or stressful conditions, and treatment of MSCs with chemicals and conditions known to stress cells (extremes of pH, high-molarity NaCl or detergents) produced similar “pseudoneuronal” morphology and increased MSC staining for neuronal markers. Strangely, pretreatment of MSCs with cycloheximide (an antibiotic that inhibits translation) failed to abrogate this response, suggesting that no new gene expression is required for cells to assume this pseudoneuronal morphology. These findings suggest that neural induction of MSCs in culture is largely an artifact (Lu, Blesch and Tuszynski 2004). Other studies have implanted MSCs into the brains of laboratory animals in the hope that a neural environment can induce neuronal differentiation in MSCs, but the implanted cells showed a spherical morphology with few extensions and connections with other cells (Zhao et al 2002).

With respect to MSC neuronal differentiation, BMSCs have definitely received the most attention. However, other types of MSCs have the capacity to form neuron-like cells (Chen, He and Zhang 2009; Chang et al 2010; Jiang et al 2010; Lim et al 2010). To date there have been few head-to-head comparisons of the efficiency of neural induction between distinct MSC populations, and this is probably a function of the variability of MSC neural induction. One study found that neural induction of UCSCs and BMSCs produces dopaminergic neurons with roughly equal efficiencies (Datta et al 2011).

Conclusion
Are BMSCs significantly different or relatively similar to MSCs from other tissue sources? The extensive research on BMSCs has provided a wealth of data that we can use for comparison with other MSCs. Work on MCSs from other tissues strongly suggests that genuine similarities exist between BMSCs and other types of MSCs. All these MSCs, with a few exceptions, display roughly the same set of cell surface proteins (De Ugarte et al 2003a; Musina, Bekchanova and Sukhikh 2005). For the most part, clonal differences in specific MSC populations notwithstanding (Zuk et al 2002; Guilak et al 2006), can differentiate into osteocytes, chondrocytes, or adipocytes (Pittenger et al 1999; Pontos et al 2006), and BMSCs and ADSCs utilize common pathways to differentiate into these distinct cell types (Liu et al 2007). They also express a common core of genes and proteins that distinguish them from other cell types.

Despite these similarities, there are also some stark differences between various MSCs from assorted tissues. First of all, the efficiencies with which these different MSC populations differentiate into osteocytes, chondrocytes, and adipocytes widely differ. Secondly, even though BMSCs and ADSCs use a set of common genes for early differentiation into all three lineages, they recruit different sets of genes for later differentiation and maturation into fully differentiated cells (Liu et al 2007; Kim and Im 2010). Thirdly, varied MSC populations differ with regards to their stemness. UCSCs share more genes in common with embryonic stem cells than BMSCs, and are, therefore, more primitive. They also express more angiogenesis and growth related genes. On the other hand, the gene expression profiles of BMSCs are much more significantly altered under different culture conditions, and express more osteogenesis genes (Hsieh et al 2010). Fourth, even though MSC populations commonly express a core set of genes(Winter et al 2003; Lee et al 2004a; Djouad et al 2005; Wagner et al 2005; Aranda et al 2009; Jansen et al 2010), gene expression profiles of distinct MSC populations differs substantially. For example, UCSCs and umbilical cord blood-derived MSCs(UBSCs) show remarkable differences in gene expression. Gene expression profiles from UBSCs revealed that genes involved in anatomical structure and multicellular organism development, osteogenesis and the immune system were expressed at high levels. However in UCSCs, genes related to cell adhesion, neurogenesis, morphogenesis, secretion and angiogenesis were more highly expressed (Secco et al 2009). Fifth, even though distinct MSC populations express very similar sets of proteins (Roche et al 2009), there are significant differences (Maurer 2011). Finally, the differentiation requirements for each MSC population differ, and these differences are a result of the signature gene expression profiles of each MSC population.

Thus, MSCs represent a familial cell type, but each distinctive MSC population represents a particular subfamily of this cell type family. While some subfamilies are clearly more closely related to some than others, these MSC subfamilies constitute the constituents that compose the MSC cell type.

AUTHOR BIO:
Michael Buratovich received his Ph.D. in Cell and Developmental Biology from UC Irvine in the laboratory of Peter Bryant where he worked on tumor suppressor genes in Drosophila melanogaster. He worked as a postdoctoral research fellow at Sussex University with Robert Whittle on the role of Wnt proteins in patterning the peripheral nervous system, and at University of Pennsylvania with Betsy Wilder on Wnt signaling during development. Since 1999, he has been a member of the faculty of Spring Arbor University (Spring Arbor, MI) in the Biochemistry department. He also served as a visiting scientist at Boston University in the laboratory of Joseph Ozer where he worked on the role of basal transcription factors in stem cell differentiation, and has collaborated with Amr Amin at the University of Al-Ain on cancer research. He is zealous about communicating science to the public and passionately blogs at http://www.beyondthedish.wordpress.com.

Michael Detamore‘s laboratory at the University of Kansas Medical Center has used mesenchymal stem cells from connective tissue in human umbilical cord tissue to form structures that have some although not all features of human hair.

Human umbilical cord contains a unique connective tissue called “Wharton’s jelly.” Wharton’s jelly expands after birth to staunch bleeding from the umbilical veins and arteries. Within Wharton’s jelly is a mesenchymal stem cell population called Wharton’s Jelly Mesenchymal Stromal Cells or WJMSCs. Experiments in a variety of labs have shown that WJMSCs have differentiate into bone, cartilage, muscle, and neural cells. Recently, several labs have used three-dimensional culture systems to get WJMSCs to differentiate into tubular and endometrial cells.

Detamore and his co-workers used this three-dimensional culture system to differentiate WJMSCs into layered structures that expressed some of the genes associated with hair follicles and also looked like follicles. The cells formed small spheres of cells known as “spheroids.” Detamore and his colleagues showed that these spheroids can form bone, but during done differentiation, Detamore and the other workers in his lab noticed that the spheroids form protrusions that looked like hairs. See for yourself below.

Patterns of hair-like structures in DWJM. Hair-like structures were observed growing out of DWJM 2 weeks following WJMSC seeding and osteogenic induction. In phase-contrast microscopy pictures (A, B) taken during culture, hair-like structures were observed to grow under the outer layer of the DWJM and in some cases either successfully protruded through the outer layer (A) or just caused a protrusion of the outer layer of DWJM (B). Also, the hair-like structures were either straight (A, C–E) or coiled (B). The green arrows in picture (A) pointed to the outer layer covering DWJM. The hair-like structure caused the outer layer covering DWJM to appear lifted up. In this DWJM, 2 hair-like structures protruded through DWJM (red arrows) seen in (A, C–E). A second hair-like structure (red arrows) protruded through the outer layer of DWJM; however, this hair-like structure was surrounded by tissue material (E). In (F), multiple areas of protrusions noted (black arrows). In (C–E) pictures, material was visualized using Nikon SMX1500 dissecting microscope and pictures were taken using Optem DC50NN camera. Scale bars represent 100 μm.

The other side of the spheroid did form bone, but the hair-like structures did eventually form bone. There are specific genes that are expressed in hair follicles, and these can be used to determine if the projections are actually hair follicles. One of these genes, cytokeratin 19 or CK19, was expressed at pretty high levels in the hair follicles. Another hair-specific gene, CK15 was also expressed in the hair-like structures.

Are these real hair follicles? Probably not, but they seem to be on their way to making hair follicles. Furthermore, the production of these hair-like structures was rather easy. If WJMSCs could be used to make hair, then they might be useful for cosmetic procedures that replace lost hair follicles as a result of baldness.